Dystonin-A3 upregulation is responsible for maintenance of tubulin acetylation in a less severe dystonia musculorum mouse model for hereditary sensory and autonomic neuropathy type VI

Abstract

Hereditary sensory and autonomic neuropathy type VI (HSAN-VI) is a recessive human disease that arises from mutations in the dystonin gene (DST; also known as Bullous pemphigoid antigen 1 gene). A milder form of HSAN-VI was recently described, resulting from loss of a single dystonin isoform (DST-A2). Similarly, mutations in the mouse dystonin gene (Dst) result in severe sensory neuropathy, dystonia musculorum (Dst^dt). Two Dst^dt alleles, Dst^dt-Tg4 and Dst^dt-27J, differ in the severity of disease. The less severe Dst^dt-Tg4 mice have disrupted expression of Dst-A1 and -A2 isoforms, while the more severe Dst^dt-27J allele affects Dst-A1, -A2 and -A3 isoforms. As dystonin is a cytoskeletal-linker protein, we evaluated microtubule network integrity within sensory neurons from Dst^dt-Tg4 and Dst^dt-27J mice. There is a significant reduction in tubulin acetylation in Dst^dt-27J indicative of microtubule instability and severe microtubule disorganization within sensory axons. However, Dst^dt-Tg4 mice have no change in tubulin acetylation, and microtubule organization was only mildly impaired. Thus, microtubule instability is not central to initiation of Dst^dt pathogenesis, though it may contribute to disease severity. Maintenance of microtubule stability in Dst^dt-Tg4 dorsal root ganglia could be attributed to an upregulation in Dst-A3 expression as a compensation for the absence of Dst-A1 and -A2 in Dst^dt-Tg4 sensory neurons. Indeed, knockdown of Dst-A3 in these neurons resulted in a decrease in tubulin acetylation. These findings shed light on the possible compensatory role of dystonin isoforms within HSAN-VI, which might explain the heterogeneity in symptoms within the reported forms of the disease.

Introduction

Dystonin (Dst), also known as bullous pemphigoid antigen 1 (Bpag1), is a massive cytoskeletal-linker protein (>600 kDa) belonging to the spectraplakin family of proteins (1–4). Differential expression of Dst isoforms occurs in neuronal, muscular and epithelial tissues. Furthermore, alternative splicing of 5′ exons yields 3 major neuronal isoforms: Dst-A1, Dst-A2 and Dst-A3 (5,6). While all three isoforms possess central plakin and spectrin repeat domains and a C-terminus microtubule binding domain (MTBD), they are unique in the organization of their N-termini. Dst-A1 possesses two calponin homology repeats that make up a functional actin-binding domain (ABD) (7,8). This allows Dst-A1 to participate in crosslinking of actin microfilaments with microtubules (9). Dst-A2 also possesses an ABD, which is preceded by a transmembrane domain (TMD) (9). This unique N-terminus localizes Dst-A2 to perinuclear membranes and surrounding microfilaments (9–11). Though it has a specific intracellular localization, DST-A2 is involved in many different cellular roles, including microtubule stability, organelle integrity and intracellular transport (12–16). Finally, Dst-A3 possesses a putative myristoylation motif (myr) followed by a single calponin homology domain, which dramatically reduces its affinity for actin (6,7,10). It is thought that myristoylation of Dst-A3 allows its localization to the plasma membrane, though its specific function remains unknown (7).

Hereditary sensory and autonomic neuropathy type VI (HSAN-VI) is the recessive human disease that results from mutations in the DST gene. The disease was first classified in 2012 when three Ashkenazi infants presented with joint contractures, dysautonomias and severe psychomotor retardation, which ultimately led to death before the age of two (17). It was discovered that these individuals possessed a homozygous frameshift mutation that disrupted the C-terminal MTBD, which is common to all three DST isoforms. More recently, two different compound heterozygous mutations have also been found to result in a non-lethal form of HSAN-VI. All three affected siblings presented with joint deformities, reduced pain and tactile sensation and an array of autonomic irregularities including hypohidrosis, chronic diarrhea and sexual dysfunction. It was determined that their novel mutations only affected the expression of DST-A2 (18), suggesting the importance of DST-A1 and DST-A3 isoforms.

Similarly, a murine disease known as dystonia musculorum (Dst^dt) also arises due to mutations in the Dst gene. Dst^dt mice first present with uncoordinated movements at postnatal day 10. Phenotype then rapidly worsens to include severe ataxia, writhing and twisting of the trunk and eventually death by about 3 weeks of age (19,20). In accordance with this, the major cell types affected in Dst^dt mice are the dorsal root ganglion (DRG) neurons involved in proprioception. They exhibit axonal swellings, a hallmark of Dst^dt, and undergo rapid neurodegeneration (11,13,21–23). Of the numerous Dst^dt alleles, Dst^dt-Tg4 and Dst^dt-27J are among the best studied. Dst^dt-27J first arose by spontaneous mutation at The Jackson Laboratory and lacks all three neuronal Dst isoforms (24,25; Fig. 1). These mice present with pronounced ataxia and severe dystonic postures, with death occurring around postnatal day 17–18. Conversely, Dst^dt-Tg4 mice arose by a transgene insertion–deletion event that disrupted expression of Dst-A1 and -A2, but left Dst-A3 intact (6,8,26; Fig. 1). They exhibit a milder phenotype and generally live a few days longer than Dst^dt-27J mice.

Figure 1

Schematic of the neuronal Dst isoforms expressed in Dst^dt-27J and Dst^dt-Tg4 mice. The more severe Dst^dt-27J mice lack expression of all three neuronal Dst isoforms. The Dst^dt-Tg4 mice possess a mutation that affects only Dst-A1 and Dst-A2 expression, with Dst-A3 remaining intact. They also have a milder disease presentation and a slightly longer lifespan.

Previous studies have described massive disorganization of the cytoskeleton in the neurons of Dst deficient mice, with the microtubule network showing substantial defects (10,14,27,28). Work done in our laboratory has also characterized a significant reduction in microtubule stability within Dst^dt-27J sensory neurons, with evidence pointing towards Dst-A2 as the major isoform responsible for this (12). Here we sought to validate these results in Dst^dt-Tg4 sensory neurons, as Dst-A2 is also lacking in these mice. Surprisingly we did not observe any obvious changes in microtubule stability in Dst^dt-Tg4 primary sensory neurons or DRGs at either the pre-phenotype or phenotype stages. Further analysis of dorsal root axons revealed cervical and thoracic level Dst^dt-Tg4 DRGs to have milder defects in microtubule organization compared to Dst^dt-27J. Although both alleles are present on different genetic backgrounds, the major difference between the two is the retention of Dst-A3 in Dst^dt-Tg4 mice. Through evaluation of Dst-A3 transcript levels in Dst^dt-Tg4 neural tissues, we determined that upregulation is most dramatic in tissues that exhibit major defects due to loss of Dst. We also observed that the retention of tubulin acetylation in Dst^dt-Tg4 sensory neurons is specifically a result of this increase in Dst-A3 transcript levels. These results are consistent with Dst-A3 taking on a compensatory role when Dst-A1 and -A2 are lost. Collectively, these findings suggest that microtubule instability is not central to initiation of Dst^dt pathogenesis though it may contribute to disease severity and that Dst-A3 upregulation compensates for the loss of other Dst isoforms.

Results

Dst^dt pathogenesis is not driven by microtubule instability

Tubulin acetylation (Ac) can be used as an indicator for microtubule stability. This post-translational modification of tubulin occurs on lysine 40 of the alpha-tubulin subunit and marks stability of polymerized microtubules as indicated by their resistance to colchicine and nocodazole treatment (29–31). Previous work on Dst^dt-27J DRGs reported a reduction in the ratio of Ac-tubulin to total alpha tubulin at the pre-phenotype stage (P4) and phenotype stage (P15) as assessed by enhanced chemiluminescence immunoblot (12). We first set out to validate these findings using more quantitative fluorescent immunoblotting techniques. We observed a significant reduction in the ratio of Ac-tubulin to alpha-tubulin in phenotype stage (P15) Dst^dt-27J DRG tissue (Fig. 2A). This reduction was further confirmed in a homogenous population of P5 primary sensory neurons grown in culture, suggesting that this pathology precedes disease onset (Fig. 2B).

Figure 2

Dst^dt-27J sensory neurons have reduced levels of Ac-tubulin. (A) Fluorescence immunoblot of Ac-tubulin and alpha-tubulin loading control for P15 Dst^dt-27J DRG tissue, quantification of Ac-tubulin:alpha-tubulin ratio shown on the right (n = 6; ^**P-value < 0.01). (B) Fluorescence immunoblot of Ac-tubulin and alpha-tubulin loading control for P5 Dst^dt-27J primary sensory neurons grown in culture, quantification of Ac-tubulin:alpha-tubulin ratio shown on the right (n = 4; ^*P-value < 0.05). All lanes in immunoblot represent DRGs from a single mouse, 10 μg of total protein per lane. Graphical data represented as mean ± SEM, statistical analysis by two-tailed Student’s t-test.

To determine whether Dst^dt-Tg4 sensory neurons also had impaired tubulin acetylation, we evaluated both pre-phenotype (P5) and phenotype (P15) stage primary sensory neurons and DRG tissue by fluorescence immunoblot. Surprisingly, regardless of in vivo or in vitro analysis at either P5 or P15 developmental stages, we did not observe any changes in the ratio of Ac-tubulin to overall alpha-tubulin between wild type (WT) and Dst^dt-Tg4 samples (Fig. 3A–D). Additionally, immunofluorescent staining of Ac-tubulin in P15 Dst^dt-Tg4 DRG tissue showed no difference in signal intensity compared to WT DRGs (Fig. 3E).

Figure 3

Tubulin-acetylation is not altered in Dst^dt-Tg4 sensory neurons. Fluorescence immunoblot analysis of Ac-tubulin and alpha-tubulin loading control for (A) P5 Dst^dt-Tg4 DRGs (n = 5), (B) P5 Dst^dt-Tg4 primary sensory neurons grown in culture (n = 3), (C) P15 Dst^dt-Tg4 DRGs (n = 5) and (D) P15 Dst^dt-Tg4 primary sensory neurons grown in culture (n = 4). Corresponding quantification for each Ac-tubulin:alpha-tubulin ratio is shown on the right of each immunoblot. All lanes of immunoblots represent DRG lysates from a single mouse, 10 μg of protein per lane. Graphical data represented as mean ± SEM, all comparisons were non-significant as assessed by two-tailed Student’s t-test (P-value > 0.05). (E) Immunofluorescence micrographs of P15 WT and Dst^dt-Tg4 DRG tissue stained positive for Ac-tubulin (green) and alpha-tubulin (red; Micrographs at 20× magnification).

As tubulin acetylation in Dst^dt-Tg4 sensory neurons is unaffected, we decided to assess a second marker for microtubule stability. Tubulin detyrosination had previously been reported to be unaltered in Dst^dt-27J DRGs (12), which we have also confirmed here in P15 Dst^dt-27J DRG tissue (Supplementary Material, Fig. S1). However, tubulin detyrosination had yet to be assessed in Dst^dt-Tg4 sensory neurons. This post-translational modification involves the proteolytic removal of the C-terminal tyrosine residue on alpha-tubulin subunits within polymerized microtubules (30,32). Using the same experimental setup as we did for tubulin acetylation, we assessed the ratio of detyrosinated-tubulin to overall alpha-tubulin. Both P5 and P15 Dst^dt-Tg4 DRGs and sensory neurons had similar detyrosinated-tubulin to alpha-tubulin ratios as WT littermates (Fig. 4A–D). Taken with the normal tubulin acetylation also found, these data suggest that Dst^dt-Tg4 sensory neurons do not have impaired microtubule stability. This is in contrast to Dst^dt-27J sensory neuron microtubules, which do exhibit reduced microtubule stability even before the onset of phenotype (12).

Figure 4

Detyrosinated-tubulin levels are not altered in Dst^dt-Tg4 sensory neurons. Fluorescence immunoblot analysis of detyrosinated-tubulin and alpha-tubulin loading control for (A) P5 Dst^dt-Tg4 DRGs (n = 5), (B) P5 Dst^dt-Tg4 primary sensory neurons grown in culture (n = 3), (C) P15 Dst^dt-Tg4 DRGs (n = 5) and (D) P15 Dst^dt-Tg4 primary sensory neurons grown in culture (n = 3). Corresponding quantification for each detyrosinated-tubulin:alpha-tubulin ratio is shown on the right of each immunoblot. All lanes of immunoblots represent DRG lysates from a single mouse, 10 μg of protein per lane. Graphical data represented as mean ± SEM, all comparisons were non-significant as assessed by two-tailed Student’s t-test (P-value > 0.05).

Dst^dt-27J and Dst^dt-Tg4 mice show disparate patterns of microtubule disorganization in dorsal root axons

As there were no significant changes in microtubule stability of Dst^dt-Tg4 DRGs, we wanted to assess if microtubule organization was also unaffected. Using electron microscopy, we assessed the cytoskeletal organization within dorsal root axons from P15 WT, Dst^dt-Tg4 and Dst^dt-27J mice (Fig. 5). As expected, Dst^dt-27J axons showed massive disorganization of the microtubule network at the cervical, thoracic and lumbar levels. This was marked by short randomly oriented microtubules (white arrows) and areas completely devoid of microtubules (black arrows). Dst^dt-Tg4 axons showed only mild defects at the cervical and thoracic levels, particularly in polarization along the length of the axon. However, their axons at the lumbar region appeared to be just as impaired as in Dst^dt-27J.

Figure 5

Microtubule organization in Dst^dt-Tg4 dorsal root axons shows milder defects compared to Dst^dt-27J. Electron micrographs of P15 WT, Dst^dt-Tg4, PrP;Dst^dt-Tg4 and Dst^dt-27J dorsal root axons from cervical, thoracic and lumbar levels. At all levels Dst^dt-27J appears to have major defects in microtubule network organization. Conversely, Dst^dt-Tg4 and PrP;Dst^dt-Tg4 dorsal root axons at the cervical and thoracic levels show much milder defects. However, their lumbar level dorsal root axons appear to have microtubule defects similar to those seen in Dst^dt-27J. Black arrows indicate areas devoid of microtubules, white arrows indicate disorganized microtubules (scale bars = 100 nm).

Considering the severe lumbar defects in microtubule organization in Dst^dt-Tg4 mice, we wanted to assess whether restoration of the Dst-A2 isoform could improve these impairments. To test this we used the PrP-Dst-A2/PrP-Dst-A2;Dst^dt-Tg4 transgenic mouse model. PrP-Dst-A2/PrP-Dst-A2;Dst^dt-Tg4 mice (herein referred to as PrP;Dst^dt-Tg4) are Dst^dt-Tg4 mice in which there is transgenic expression of full-length Dst-A2 cDNA under the neuronal prion protein promoter (PrP) (11). Interestingly, we did not observe any major differences between PrP;Dst^dt-Tg4 and Dst^dt-Tg4 axons (Fig. 5). The lack of any discernable difference could be due to the fact that Dst-A1 is still absent or possibly due to the low levels of the Dst-A2 transgene expressed in DRG tissue (11). It may even be that differential expression of the transgene in DRGs along the length of the spinal cord contributes to the lumbar defects that persist in PrP;Dst^dt-Tg4 mice. In order to address this concern we evaluated Dst-A2 expression within cervical, thoracic and lumbar level DRGs from WT and PrP;Dst^dt-Tg4 mice (Supplementary Material, Fig. S2). We however did not see a significant difference in transgene expression at any of the spinal levels from PrP;Dst^dt-Tg4 mice, suggesting that Dst-A2 is uniformly expressed in DRGs at all levels.

Curiously, the observed patterns of microtubule disorganization mirror the way in which both Dst^dt-Tg4 and Dst^dt-27J mice present their phenotype. By P15, Dst^dt-27J mice have lost all forelimb and hindlimb movement coordination and exhibit dystonic postures (Fig. 6A left panel), in accordance with defects observed along the full length of the spinal cord. Conversely Dst^dt-Tg4 mice are not as severely affected at this age as they can still support themselves and coordinate movement with their forelimbs and do not adopt dystonic postures. However, in parallel with lumbar spinal cord defects, their hindlimbs do exhibit impaired movement coordination (Fig. 6A right panel). PrP;Dst^dt-Tg4 mice also exhibit a phenotype similar to Dst^dt-Tg4, although onset is delayed. They retain forelimb control, but hindlimb coordination is impaired, in line with the impairments in microtubule disorganization observed.

Figure 6

Dst^dt-Tg4 mice have a milder phenotype than Dst^dt-27J. (A) Phenotype stage (P15) Dst^dt-27J mouse (left) and Dst^dt-Tg4 mouse (right). Dst^dt-27J has lost all limb coordination and shows writhing and twisting of the trunk. The Dst^dt-Tg4 mouse does not have dystonic postures. They can support themselves and direct movement with their forelimbs, however their hindlimbs often exhibit poor coordination and drag behind them when they walk. (B–C) Results from the horizontal ladder test shows that forelimb and hindlimb coordination simultaneously decline early (P10–12) in Dst^dt-27J mice (n = 7 for WT, and n = 3 for Dst^dt-27J though only one mouse survived to P18 time point). Conversely, Dst^dt-Tg4 mice exhibit a milder impairment in forelimb coordination compared to Dst^dt-27J. Hindlimb defects are also less severe compared to Dst^dt-27J. The frequency of detectable hindlimb faults appears to gradually increase over time in Dst^dt-Tg4 mice (n = 3, though for Dst^dt-Tg4 only one mouse survived to P20 time point). It should however be noted that hindlimb dragging, a common Dst^dt-Tg4 phenotype, could not be measured by this test (^*P-value < 0.05, ^**P-value < 0.005, ^***P-value < 0.0005 comparison between Dst^dt-27J and Dst^dt-Tg4; # P-value < 0.05, ## P-value < 0.005 comparison between Dst^dt-27J and their WT littermates; $ P-value < 0.05, $$ P-value < 0.005 comparison between Dst^dt-Tg4 and their WT littermates).

Results from horizontal ladder test confirm that Dst^dt-27J forelimb and hindlimb coordination is more severely affected compared to WT littermates and Dst^dt-Tg4 mice. Defects are observed as early as P10, and coordination of both limb pairs continues to worsen until death (Fig. 6B and C). Dst^dt-Tg4 mice show some impairment in forelimb coordination, though this level of impairment is maintained at a relatively steady state from P10 until death (Fig. 6B). Dst^dt-Tg4 hindlimb coordination however appears to gradually decline over the testing period (Fig. 6C). Although no significant difference was detectable between Dst^dt-Tg4 forelimb and hindlimb coordination by horizontal ladder test, qualitative assessment of these mice suggests a greater impairment in hindlimb function and coordination. As Dst^dt-Tg4 mice tend to drag their hindlimbs behind them when they walk, the severity of their hindlimb defect would not be detectable by this test.

Figure 7

Late stage Dst^dt-Tg4 DRGs show no further changes to microtubule network organization. (A) Fluorescence immunoblot analysis of Ac-tubulin and alpha-tubulin loading control for P21 Dst^dt-Tg4 DRGs, quantification of Ac-tubulin:alpha-tubulin ratio shown on the right (n = 3, ns P-value > 0.05). (B) Fluorescence immunoblot analysis of detyrosinated-tubulin and alpha-tubulin loading control for P21 Dst^dt-Tg4 DRGs, quantification of detyrosinated-tubulin:alpha-tubulin ratio shown on the right (n = 3, ns P-value > 0.05). (C) Electron micrographs of P20 WT, Dst^dt-Tg4 and PrP;Dst^dt-Tg4 dorsal root axons, and P40 PrP;Dst^dt-Tg4 dorsal root axons from cervical, thoracic and lumbar levels. No further changes are observed from P15 electron micrographs. Black arrows indicate areas devoid of microtubules, white arrows indicate disorganized microtubules (scale bars = 100 nm).

Microtubule stability and organization remain unchanged in late-stage Dst^dt-Tg4 DRGs

Considering Dst^dt-Tg4 mice live longer than Dst^dt-27J (Fig. 8L), we wanted to make certain that we were not missing any microtubule-related defects at P15, which is not necessarily ‘end-stage’ for Dst^dt-Tg4 mice. As such, ratios of Ac-tubulin and detyrosinated-tubulin to overall alpha-tubulin were determined for P21 Dst^dt-Tg4 and WT DRGs (Fig. 7A and B). There were no significant differences observed for either condition. We also assessed microtubule organization in cervical, thoracic and lumbar dorsal root axons from P20 Dst^dt-Tg4 and PrP;Dst^dt-Tg4 mice, as well as from P40 PrP;Dst^dt-Tg4 mice (Fig. 7C). Cervical and thoracic level microtubule organization in Dst^dt-Tg4 and PrP;Dst^dt-Tg4 axons was similar to that at P15. Lumbar dorsal root axons continued to display the same level of cytoskeletal disorganization. No further defects were observed in late-stage (P40) PrP;Dst^dt-Tg4 dorsal root axons. From this we find that 1) microtubule stability is unaltered in Dst^dt-Tg4 DRGs and 2) defects in microtubule organization are present, particularly at the lumbar level, which correlates with presentation of phenotype.

In order to address whether the observed differences between Dst^dt-27J and Dst^dt-Tg4 mice could be due to differences in cell death, we evaluated apoptosis by TUNEL labeling, along with a cell death analysis by assessment of histological features via hematoxylin and eosin staining in phenotype stage DRGs (Supplementary Material, Fig. S3). We did not observe any overt difference in number of TUNEL positive sensory neurons between Dst^dt-27J and Dst^dt-Tg4 at any spinal level. We do however see that there is an abundance of TUNEL positive glial cells in Dst^dt-27J and Dst^dt-Tg4 sections, these can primarily be seen at cervical and thoracic levels. Histological analysis reveals that there may be fewer sensory neurons present within Dst^dt-27J when compared to Dst^dt-Tg4 mice, however this may be due to the somewhat smaller size of the mice and their DRGs. Further quantitative analysis of sensory neuron number over multiple time points is necessary to determine whether this is a result of cell death or other factors.

Dst-A3 is upregulated in Dst^dt-Tg4 DRGs and spinal cord

As we were surprised to find that microtubule stability was unimpaired in Dst^dt-Tg4 sensory neurons, we considered possibilities for why this pathology is not shared by the different Dst alleles. Although Dst^dt-27J and Dst^dt-Tg4 mice are on different genetic backgrounds, C57BL6 and CD1, respectively, the major difference between them is that Dst^dt-Tg4 retains Dst-A3 expression. Perhaps this isoform has a role in maintaining microtubule stability when the other neuronal isoforms are lost. To evaluate this, we assessed Dst-A3 gene expression in WT and Dst^dt-Tg4 DRGs by real-time quantitative reverse transcription-polymerase chain reaction (RT-qPCR). Interestingly, we observed an ∼4-fold increase in Dst-A3 transcript levels in P5 Dst^dt-Tg4 DRGs (Fig. 8A), while P15 Dst^dt-Tg4 DRGs showed an ∼3-fold increase in transcript levels (Fig. 8B). This pattern of upregulation suggests that Dst-A3 may have a compensatory role.

Figure 8

Dst-A3 is upregulated in Dst^dt-Tg4 dorsal root ganglia and spinal cord tissue. Quantification of the relative levels of Dst-A3 mRNA in P5 (A) and P15 (B) WT and Dst^dt-Tg4 DRGs (n = 4, ^*P-value < 0.05, ^**P-value < 0.01). Relative Dst-A3 expression levels also assessed in P15 WT, heterozygous Dst^dt-Tg4/+, Dst^dt-Tg4 and PrP;Dst^dt-Tg4 DRGs by RT-qPCR (C) and RT-PCR (D) (n = 3, ^*P-value < 0.05). Dst-A3 mRNA expression also assessed in P15 WT and Dst^dt-Tg4 by RT-qPCR and RT-PCR in spinal cord (E,F) and cerebral cortex (G,H) tissues (n = 3, ^*P-value < 0.05). (I, J)Dst-A3 expression profile assessed in P15 WT cerebral cortex, spinal cord and DRGs by RT-qPCR and RT-PCR (n = 3, ^**P-value < 0.01). Graphical data represented as mean ± SEM, statistical analysis by two-tailed Student’s t-test or one-way ANOVA and Tukey’s post hoc test, where appropriate. Actin (actb) used for loading control in RT-PCR analysis, while Ppia and Rps18 used to normalize in RT-qPCR experiments. (K) Quantification of Dst-A3 mRNA levels in P15 WT and Dst^dt-Tg4 DRGs separated by spinal level (cervical, thoracic and lumbar). No significant differences observed within DRGs from each genotype (n = 3, ns P-value > 0.05). Dst-A3 expression normalized to Ppia and Rps18. Data represented as mean ± SEM, statistical analysis by one-way ANOVA and Tukey’s post hoc test. (L) Kaplan–Meier survival curve indicates that Dst^dt-Tg4 mice, including those on the mixed CD1/C57BL6 background, live longer than Dst^dt-27J mice.

To rule out the possibility that the inserted hsp68-LacZ transgene of the Dst^dt-Tg4 allele is responsible for Dst-A3 upregulation, we assessed Dst-A3 transcript levels in P15 heterozygous Dst^dt-Tg4/+ DRGs and in P15 PrP;Dst^dt-Tg4 DRGs. As Dst^dt-Tg4/+ DRGs have a 50% reduction in Dst-A1 and -A2 expression, and yet Dst-A3 transcript levels remain unchanged from WT, this indicates that the hsp68-LacZ transgene alone is not capable of increasing Dst-A3 expression (Fig. 8C and D). PrP;Dst^dt-Tg4 DRGs on the other hand continue to overexpress Dst-A3 at a level similar to Dst^dt-Tg4 DRGs (Fig. 8C and D). This could be because Dst-A2 levels in PrP;Dst^dt-Tg4 DRGs are not restored to WT levels (11) or perhaps Dst-A3 is compensating for a role normally performed by Dst-A1.

We also examined Dst-A3 transcript levels in other neuronal tissues, such as the spinal cord and cerebral cortex, to determine whether this was simply a global effect driven by loss of Dst-A1 and -A2. Dst^dt-Tg4 spinal cord exhibited only an ∼50% increase in Dst-A3 gene expression compared to WT (Fig. 8E and F), while Dst^dt-Tg4 cortex did not show any significant difference (Fig. 8G and H). Interestingly, when we collectively look at the Dst-A3 patterns of expression in Dst^dt-Tg4 tissues, it appears to occur in a manner that reflects the necessity of Dst for that specific tissue, i.e. Dst-A3 overexpression is greatest in DRGs, the tissue where loss of Dst has the most severe impact. Milder pathologies are also observed in the spinal cord, where we find Dst-A3 to be moderately upregulated, and no changes are present in the cerebral cortex where loss of Dst has no significant impact. These results are consistent with Dst-A3 taking on a compensatory role in tissues where Dst is needed, when Dst-A1 and -A2 are lost.

To put this in perspective, we assessed the relative levels of Dst-A3 expression normally present in P15 WT neural tissues (Fig. 8I and J). Unexpectedly, highest levels were observed in the spinal cord, being roughly 12-fold higher than in cerebral cortex or DRGs. This might be indicative of a unique/novel role for Dst-A3 in the spinal cord. In any case, these results help us to gauge the impact of a 50% increase in spinal cord, and a 3–4-fold increase in DRGs as it relates to Dst-A3 expression in Dst^dt-Tg4 mice. Furthermore, evaluation of the three neuronal Dst isoforms in P15 WT DRGs reveals that Dst-A3 is expressed at extremely low levels compared to both Dst-A1 and -A2 (Supplementary Material, Fig. S4A and B). This suggests that Dst-A3 may typically not be as important as the other two isoforms in DRGs. However, upon the loss of Dst-A1 and -A2, a 3–4-fold upregulation in Dst-A3 may have a significant biological effect, further suggesting a compensatory role.

We then examined the Dst-A3 expression profile for WT and Dst^dt-Tg4 DRGs by spinal level to determine whether Dst-A3 has a role in maintaining microtubule organization. We observed no significant differences in Dst-A3 expression between cervical, thoracic or lumbar Dst^dt-Tg4 DRGs, indicating no correlation between level of Dst-A3 expression and degree of microtubule disorganization (Fig. 8K). Further examination of microtubule stability in cervical, thoracic and lumbar DRGs also revealed no overt changes in the tubulin acetylation or detyrosination status of Dst^dt-Tg4 DRGs compared to WT (Supplementary Material, Fig. S5A and B).

As both Dst^dt-Tg4 and Dst^dt-27J alleles are on different genetic backgrounds (CD1 and C57BL6, respectively) we wanted to rule out the possibility that genetic factors other than Dst could be responsible for the differences between these two alleles. For this we assessed survival of Dst^dt mice, including Dst^dt-Tg4 mice on a mixed CD1/C57BL6 background (Fig. 8L). Since these mixed background Dst^dt-Tg4 mice have similar survival to Dst^dt-Tg4 on CD1 background, it is likely that the genetic background doesn’t have a major effect on disease presentation.

Figure 9

Knockdown of Dst-A3 in Dst^dt-Tg4 primary sensory neurons results in loss of tubulin acetylation. (A) Quantification of the relative levels of Dst-A3 mRNA in Dst^dt-Tg4 and WT sensory neurons post adenoviral transduction with either Dst shRNA or scrambled control. Dst^dt-Tg4 sensory neurons show a significant reduction in Dst-A3 mRNA expression after Dst shRNA treatment compared to the scrambled control-treated sensory neurons. Dst-A3 expression normalized to Ppia and Rps18. Data represented as mean ± SEM, statistical analysis by one-way ANOVA and Tukey’s post hoc test (n = 3, ^*P-value < 0.05, ^**P-value < 0.01). (B) Immunofluorescence micrographs of Dst^dt-Tg4 sensory neurons post transduction by AdV-shRNA(Scrambled)-GFP (left) or by AdV-shRNA(Dst)-GFP (right). Transduced cells expressing GFP and the scrambled control are observed to have high levels of acetylated tubulin within the neuronal cell body (indicated by an asterisk). In transduced neurons expressing GFP and Dst shRNA, we observe a reduction in acetylated-tubulin levels. We also observe a non-transduced Dst^dt-Tg4 sensory neuron (GFP-negative) among the Dst shRNA group that continues to maintain a high degree of tubulin acetylation within the cell body (indicated by white arrow). Micrographs shown at 40× magnification.

Dst knockdown in Dst^dt-Tg4 sensory neurons results in loss of tubulin acetylation

Thus far we have shown that Dst-A3 is upregulated in Dst^dt-Tg4 sensory neurons, in a manner that suggests that this isoform has a compensatory role when Dst-A1 and -A2 are lost. To test whether the increase in Dst-A3 expression within Dst^dt-Tg4 sensory neurons is directly responsible for the preservation of microtubule stability, we assessed tubulin acetylation status after Dst knockdown in Dst^dt-Tg4 primary sensory neuron cultures. Using an adenovirus delivery system, shRNA specific for either the Dst mRNA or a scrambled control were introduced to sensory neuron cultures established from P15 mice. Through RT-qPCR analysis, we were able to validate that the increase in Dst-A3 expression is specific to the Dst^dt-Tg4 sensory neuron, as we observed a significant difference between WT and Dst^dt-Tg4 scrambled groups (Fig. 9A). In Dst^dt-Tg4 sensory neurons treated with Dst shRNA, Dst-A3 was knocked down by 80% relative to scrambled control (Fig. 9A). Interestingly, the Dst-A3 knockdown level achieved was comparable to normal physiological levels in WT sensory neurons, thus simply mimicking a loss of the characteristic Dst-A3 overexpression seen in Dst^dt-Tg4 DRGs.

To assess changes in microtubule stability with the loss of Dst-A3 upregulation, we proceeded with immunofluorescence analysis of acetylated-tubulin within Dst^dt-Tg4 transduced (GFP-positive) neurons. Comparison of the Dst shRNA versus scrambled control-treated Dst^dt-Tg4 sensory neurons revealed a stark difference in levels of tubulin acetylation within the cell bodies. While scrambled control-treated Dst^dt-Tg4 neurons maintained high levels of acetylated tubulin throughout the cell body cytoplasm (Fig. 9B left), the Dst shRNA-treated Dst^dt-Tg4 neurons had little or no tubulin acetylation throughout the neuron soma (Fig. 9B right). We also observed neurons in the non-transduced (GFP-negative) Dst shRNA-treated Dst^dt-Tg4 group that retained a high level of acetylated-tubulin in the soma (Fig. 9B right, white arrow). Since Dst^dt-Tg4 neurons do not express Dst-A1 or -A2, our results indicate Dst-A3 upregulation is required to achieve the improved microtubule stability observed in Dst^dt-Tg4 sensory neurons, since returning the expression to normal physiological levels is sufficient to promote a loss of stability when the other Dst isoforms are not present.

Discussion

As Dst-A2 was thought to be the major isoform responsible for microtubule stability, we expected to find defects in tubulin acetylation in Dst^dt-Tg4 DRGs, as had previously been found in Dst^dt-27J (12). Surprisingly, tubulin acetylation was unaltered in Dst^dt-Tg4 DRGs, and the organization of the microtubule network was not as severely impaired as Dst^dt-27J. Our results show that microtubule instability is not causative of the Dst^dt disease, though it may contribute to the severity. We also report an upregulation in Dst-A3 within Dst^dt-Tg4 DRGs, spinal cord tissues and sensory neurons in culture. We have determined that the increase in expression within sensory neurons is part of a compensatory mechanism when other neuronal Dst isoforms are lost. Though Dst-A3 may not typically have a major role in microtubule stability, its increased expression in sensory neurons is responsible for maintaining tubulin acetylation. This is likely one of the major reasons why we see milder pathologies and a longer lifespan in Dst^dt-Tg4 mice compared to Dst^dt-27J mice.

Differences between the Dst^dt-Tg4 and Dst^dt-27J alleles

In studying Dst, we often attempt to corroborate our findings from one Dst^dt allele in another Dst^dt allele (15). The evidence from the present study, however, shows that this may not always be possible. For the first time, we report differences in pathology between Dst^dt-Tg4 and Dst^dt-27J alleles, even though they both still succumb to the disease at an early age.

It had been previously thought that microtubule instability was a common pathology of Dst^dt, primarily due to the loss of Dst-A2 expression (12). Decreased tubulin acetylation had been observed by immunoblot analysis in Dst^dt-27J DRGs and by immunofluorescence staining of Dst^dt-27J primary sensory neurons. Through siRNA knockdown of Dst-A1 and Dst-A2 isoforms in HEK293T immortalized cells, it was found that loss of Dst-A2 resulted in the most profound defects in microtubule stability and organization (12). Based on these results, it was believed that Dst-A2 was the major isoform involved in maintaining microtubule stability; and since Dst^dt-Tg4 also lacked this isoform, we presumed microtubule stability would also be impaired. However, what was not accounted for in these studies was the impact of Dst-A3 expression on microtubules, and siRNA knockdown of this isoform was not attempted in HEK293T cells. Recent work in our laboratory has determined that HEK293T cells do not express the Dst-A3 isoform (data not shown). Thus, when Dst-A2 was knocked down, Dst-A3 would not have been able to compensate, resulting in a dramatic loss of tubulin acetylation.

We saw no change in Dst^dt-Tg4 DRG microtubule stability, which is the result of not only Dst-A3 retention, but of a compensatory effect mediated by Dst-A3 upregulation (Fig. 3 and 4). This maintenance of microtubule stability may also help to explain why defects in axonal trafficking were not previously observed in Dst^dt-Tg4 primary sensory neurons (33). Live imaging of mitochondrial movement over a very short time frame was conducted on WT and Dst^dt-Tg4 sensory neurons grown in culture. It was determined that there was no difference in number of motile mitochondria, or in their velocity along the axon. Since molecular motors such as kinesin and dynein preferentially bind and move along acetylated microtubules, this may be why no major defects in mitochondria movement were observed in Dst^dt-Tg4 sensory neurons (34–36). It also appears that the previous work describing microtubule disorganization of Dst^dt-Tg4 DRGs had only assessed DRGs of the lumbar spinal level (21). This result is in line with what we have observed for lumbar Dst^dt-Tg4 DRGs, however no previous insight had been given as to the state of microtubule organization at higher spinal cord levels. We have now determined that microtubule organization is only mildly impaired at cervical and thoracic level Dst^dt-Tg4 dorsal root axons, while major defects are present at the lumbar level (Fig. 5). As this correlates with the Dst^dt-Tg4 phenotype, we suspect that microtubule disorganization plays some role in phenotype development. It may be that Dst-A3 overexpression protects against the rapid loss of axonal cytoskeleton disorganization, but since lumbar DRGs continue to overexpress Dst-A3 and still succumb to extensive microtubule disorganization (Fig. 8K), Dst-A3 alone is clearly not enough to rescue it. Lumbar DRGs may be more vulnerable to defects since they possess the longest axons (37), or it might be due to another factor that has yet to be elucidated.

It should also be noted that the majority of studies that have reported defective axonal trafficking in Dst^dt neurons have done so using the Dst^dt-27J or Dst^tm1EFu (Dst knockout) alleles, which both lack all Dst isoforms (12,14,27,38–40). So perhaps it is a combination of microtubule instability and disorganization that contributes to impaired axonal transport. Although this is the first time we have directly compared Dst^dt-Tg4 and Dst^dt-27J mice to discover differences in their pathologies, it is possible that more differences exist between the two alleles. Identification of these would only stand to help us understand the differences in phenotype and lifespan, and also elucidate a common mechanism of death.

Insights into the role of Dst-A3

Of the three neuronal Dst isoforms, Dst-A3 is the one we know the least about. There is only one previous study that had directly evaluated Dst-A3 (also named BPAG1a3) intracellular localization by use of Flag-tagged constructs. They reported that the Dst-A3 N-terminus contains an myr, which allows this isoform to primarily localize to the plasma membrane (7). However, a prior study had been conducted on the BPAG1n3 isoform (the less abundant intermediate filament binding isoform) also using Flag-tagged constructs containing the N-terminal of this isoform (10). Since both BPAG1n3 and BPAG1a3 (Dst-A3) possess the same N-terminal sequences and only vary towards the C-terminus (4,6,41), we believe the use of N-terminal BPAG1n3 constructs would likely yield results most relevant to Dst-A3, as it is the more predominantly expressed isoform. Their study had found that the BPAG1n3/Dst-A3 N-terminal constructs localized mainly to microtubules, and this was further validated using N-terminal specific BAPG1n3/Dst-A3 antibodies. The proposed function of this isoform on microtubules was that it conferred stability, since this interaction made microtubules more resistant to nocodazole and colchicine treatment (10).

Our results further support Dst-A3’s involvement in microtubule stability since the knockdown of Dst-A3 in Dst^dt-Tg4 sensory neurons led to a loss of tubulin acetylation. However, this role may only be relevant when Dst-A3 is upregulated, as is the case for Dst^dt-Tg4 sensory neurons and cells transfected with BPAG1n3/Dst-A3 constructs. Under normal conditions Dst-A2 is most likely the major isoform responsible for microtubule stability (12), especially as Dst-A2 is also the isoform most abundantly expressed in DRGs (Supplementary Material, Fig. S4A and B).

Aside from its N-terminus, Dst-A3 contains the same central plakin domain and C-terminal Gas2-related domain as Dst-A1 and -A2. The plakin domain of Dst has previously been shown to interact with the microtubule stabilizing protein MAP1B (42). More specifically, MAP1B interaction with Dst-A2 has been shown to maintain tubulin acetylation (12). It could be that when Dst-A2 is lost, MAP1B preferentially binds to the upregulated Dst-A3 and preserves microtubule stability. Some degree of microtubule stability may also come from direct binding of Dst-A3’s Gas2-related domain with microtubules, which has been shown to protect against depolymerization (43).

Ultimately, overexpression of Dst-A3 is not enough to rescue the defects from loss of Dst-A1 and -A2. Therefore this isoform must have its own unique function within the cell and is not fully redundant to either Dst-A1 or -A2. It appears that Dst-A3 is involved in maintaining microtubule stability, however this may not be its predominant function under normal conditions. It would certainly be interesting to further investigate what role Dst-A3 plays within neurons, and as spinal cord seems to express the highest levels of this isoform (Fig. 8I and J), this tissue would be a good starting point for elucidating its quintessential function within neurons.

Dst isoform compensation in HSAN-VI?

It was previously believed that HSAN-VI was a lethal disorder of childhood (17), however the discovery of novel compound heterozygous mutations in an Italian family in 2017 has proven otherwise (18). As these patients are ∼40 years of age and were found to only lack the Dst-A2 isoform, we now know that retention of all three neuronal isoforms is not critical to survival. Considering this, we may expect for HSAN-VI to present in a variety of manners, varying in severity depending on the mutation and the isoforms affected. Our findings of Dst-A3 upregulation in Dst^dt-Tg4 sensory neurons also lead us to believe that this same mechanism of Dst isoform upregulation is possible in HSAN-VI. This may even be one of the reasons for why a much milder disease presentation is observed in the Italian patients; Dst-A3 and maybe even Dst-A1 might be expressed at much higher levels than normal in attempt to offset and compensate for the lack of Dst-A2. This concept is highly exciting as it could open up a number of different therapeutic options, aimed at altering isoform expression levels opposed to the more challenging route of gene therapy.

Conclusions

Here we have determined that the different Dst alleles do not always have identical pathologies. We have observed that microtubule instability is not causative of Dst^dt pathology, however it may be one of the factors that influences how severely the disease presents. Additionally, we have also identified an upregulation of Dst-A3 within Dst^dt-Tg4 sensory neurons, indicative of the Dst-A3 isoform taking on a compensatory role in maintenance of microtubule stability when the other neuronal isoforms are lost.

Taken together, the work presented here sheds light onto the phenotypic differences observed between different Dst alleles and the underlying genetics that cause them. This is highly relevant to human disease, as DST mutations resulting in HSAN-VI lead to a spectrum of symptoms and longevity. Prior to the discovery of the most recent set of HSAN-VI cases, it was believed that HSAN-VI was lethal before the age of two (17); however the more recent patients deficient in only DST-A2 continue to live well into their forties (18). Given the findings we report here, we might even expect to find that DST-A3 (and even possibly DST-A1) are upregulated in their neuronal tissues, potentially compensating for the normal roles of DST-A2 resulting in milder symptoms and prolonged lifespan. As such, further investigation into the possible compensatory effects of Dst isoforms will be invaluable to our understanding of HSAN-VI and in developing therapies for those affected.

Materials and Methods

Ethics statement

All experimental protocols on mice were approved by the Animal Care Committee of the University of Ottawa. Care and use of experimental mice followed the guidelines of the Canadian Council on Animal Care and the Animals for Research Act.

Animals

The Dst^dt-Tg4 mouse line arose from a transgene insertion–deletion that affected the 5′ region encoding the ABD specific to DST-A1 and DST-A2 isoforms (8,24,26). For these experiments, all Dst^dt-Tg4 mice and their WT littermates were maintained on a CD1 background (except where specifically indicated that they are on a mixed CD1/C57BL6 background). The Dst^dt-27J line arose due to spontaneous mutation and was found to express very low levels of the Dst transcript (24). These mice, along with their WT littermates, are on a C57BL6 genetic background. Genotypes for both lines of mice were determined by PCR amplification of genomic tail DNA, as described in (44).

The PrP-Dst-A2/PrP-Dst-A2;Dst^dt-Tg4 rescue mouse (referred to as PrP;Dst^dt-Tg4 mice in the text) is a transgenic mouse which exogenously expresses Dst-A2 under the control of the neuronal PrP on the Dst^dt-Tg4 background. These mice are maintained and bred on a mixed CD1/C57BL6 background. The derivation and characterization of these mice have been previously described in (11,44).

Primary sensory neuron culture and adenovirus transduction

DRGs from P5 and P15 mice were collected following the protocol previously described in (44). The individual 1.5 ml microfuge tubes containing DRGs in 1X Hank’s Balanced Salt Solution (HBSS; Gibco) were centrifuged at 300 g for 5 min. The HBSS was removed and replaced with 300 μl of prewarmed 1.7 mg/ml papain (Worthington). Tubes were then incubated in a 37°C water bath for 10 min and centrifuged at 300 g for 5 min. The papain solution was then removed and replaced with 300 μl of 2 mg/ml Collagenase A (Roche). Tubes were once again incubated in a 37°C water bath for 10 min and centrifuged at 300 g for 5 min. After centrifugation, Collagenase A was carefully removed and replaced with 1 ml DRG media (10% fetal bovine serum [FBS; Gibco], 1% penicillin/streptomycin [Gibco], in a Dulbecco’s Modified Eagle Medium [DMEM; Wisent] base). Tubes were centrifuged again at 300 g for 5 min. After centrifugation, the 1 ml of DRG media was removed and replaced with 1.3 ml fresh DRG media. Flame-polished glass Pasteur pipettes were then used to dissociate the DRG neurons by trituration. Tubes were centrifuged for a final time at 300 g for 5 min. Supernatant was removed, and the pellets were resuspended in 1 ml of fresh DRG media. Total DRG neurons from a single mouse were then seeded in a single well of a 6-well plate coated with 4% Laminin (LN2; Milipore). Plates were then transferred to a 37°C tissue culture incubator at 8.5% CO₂. Twelve hours later, full media change was performed by replacing DRG media with neuronal maintenance media (DMEM base [Wisent], 0.5% FBS [Gibco], 1% penicillin/streptomycin [Gibco], 2% B27 supplement [Gibco], 1% GlutaMax [Gibco], 16 mg/ml putrescine [Sigma-Aldrich], 400 mg/ml thyroxine [Sigma-Aldrich], 400 mg/ml triiodothyronine [Sigma-Aldrich], 6.2 ng/ml progesterone [Sigma-Aldrich], 5 ng/ml sodium selenite [Sigma-Aldrich], 100 mg/ml bovine serum albumin [BSA; Sigma-Aldrich], 5 mg/ml bovine insulin [Sigma-Aldrich], 50 mg/ml holo-transferrin [Sigma-Aldrich] supplemented with 1 mm 5-fluoro-2′-deoxyuridine [Sigma-Aldrich]). A two-thirds media change was performed every other day until protein or RNA collection on day five.

For adenovirus transduction experiments, P15 primary sensory neuron cultures were grown as usual until day five and then transduced with either custom-designed Ad-U6-shRNA(mDST)-CMV-GFP (SignaGen Laboratories) or a scrambled control Ad-U6-shRNA(Scramble)-CMV-GFP (SignaGen Laboratories) at a multiplicity of infection of 450 for both. Transduced sensory neuron cultures were maintained for 72 h, RNA was then collected or coverslips were fixed with 3% paraformaldehyde (PFA) for immunofluorescence analysis.

Immunoblotting

Protein was extracted from primary sensory neuron cultures, and from P5, P15 and P21 DRG tissue by gently homogenizing the samples with 1X RIPA buffer (Sigma-Aldrich) supplemented with 6 μl/ml phenyl methyl sulfonyl fluoride (Sigma-Aldrich). Samples were left on ice for 10 min before centrifugation at 4°C for 10 min at 12 000 g. Supernatant was then collected, and concentrations were determined by bicinchoninic acid assay (BCA assay; Pierce). Protein samples (10 μg, except where indicated otherwise) were separated by SDS-PAGE in 10% gels. PVDF membranes were incubated overnight at 4°C in a dilution of Odyssey® blocking buffer (LI-COR) and primary antibody. The following primary antibodies were used: mouse monoclonal anti-acetylated tubulin antibody (1:40 000; Sigma-Aldrich #T7451), rabbit polyclonal anti-detyrosinated tubulin antibody (1:25 000; Millipore #AB3201), mouse monoclonal anti-alpha tubulin antibody (1:50 000; Calbiochem CP06) and rabbit polyclonal anti-alpha tubulin antibody (1:20 000; Abcam ab18251). Membranes were then washed 3 times with 1X Tris-Buffered Saline and incubated for 1 h at RT in an Odyssey® blocking buffer (LI-COR) and secondary antibody solution. Infrared dye conjugated (IRDye®) secondary antibodies used include goat anti-rabbit 800CW (IgG, 1:10,000; LI-COR; 926–32211), goat anti-rabbit 680RD (IgG, 1:10,000; LI-COR; 926–68071), goat anti-mouse 800CW (IgG, 1:10,000; LI-COR; 926–32210) and goat anti-mouse 680RD (IgG, 1:10,000; LI-COR; 926–68070). Bands were visualized and quantified using the Odyssey® CLx infrared imaging system (LI-COR Biosciences).

Immunohistochemistry

Whole DRGs from P15 WT and Dst^dt-Tg4 mice were extracted following euthanasia and placed in Cryomolds® (Tissue-Tek®) containing O.C.T. compound (Tissue-Tek®) and snap frozen in liquid nitrogen. Sections of 7 μm thickness were made using a cryostat and mounted on glass slides for staining. Slides were then fixed in 4% PFA for 10 min and washed 3 times in 1X phosphate buffered saline (PBS), pH 7.4. Sections were permeabilized in 0.2% Triton-X (Sigma-Aldrich) in 1X PBS for 10 min and then blocked in 1% BSA (Sigma-Aldrich) in 1X PBS for 15 min. Primary antibody dilutions of anti-Ac tubulin antibody (1:1000; Sigma-Aldrich #T7451) and anti-alpha-tubulin antibody (1:1200; Millipore 04–1117) were applied to slides and incubated for 1 h at room temperature. Secondary antibodies Alexa Fluor 488 goat anti-mouse (1:1000; Invitrogen A-11001) and Alexa Fluor 555 goat anti-rabbit (1:1000; Invitrogen A-21428) applied to slides for 45 min at room temperature. Finally, DAPI (1:1000) was applied for 5 min at room temperature, and slides were then mounted in Dako fluorescence mounting medium (Dako S3023).

Immunocytochemistry

Transduced P15 primary sensory neuron cultures were fixed on coverslips in 3% PFA for 10 min, and subsequently washed 3 times with 1X PBS for 5 min. Cells were then permeabilized with 0.1% Triton-X (Sigma-Aldrich) in 1X PBS for 10 min, washed 3 times with 1X PBS and blocked for 1 h with 10% goat serum in 1X PBS. Primary antibody dilutions of anti-Ac-tubulin antibody (1:5000; Sigma-Aldrich #T7451) and anti-alpha-tubulin antibody (1:5000; Abcam ab18251) were made in 10% goat serum and left on coverslips overnight at 4°C. Coverslips were then washed 3 times in 1X PBS for 5 min and incubated with secondary antibody solution for 1 h at room temperature. The secondary antibody dilution consisted of Alexa Fluor 555 goat anti-mouse (1:200, Invitrogen A28180) and Alexa Fluor 647 goat anti-rabbit (1:200, Invitrogen A-21244). Coverslips were then counterstained with Hoechst (1:5000) and mounted in Dako fluorescence mounting medium (Dako S3023). Analysis was performed using a Zeiss LSM 510 Confocal Microscope equipped with Zen 2009 imaging software (Zeiss). An n = 3 was analyzed for WT and Dst^dt-Tg4 sensory neurons transduced with either shRNA or scrambled control. Images were taken at a magnification of 40×.

TUNEL labeling

Cervical, thoracic and lumbar DRGs were dissected from P15 WT, Dst^dt-27J and Dst^dt-Tg4 mice. Sections were made at a thickness of 10 μm and washed in 1X PBS. Sections were fixed for 20 min in 4% PFA, followed by a 30 min wash in 1X PBS. Permeabilization was carried out using ice cold 0.1% sodium citrate/0.1% Triton X-100 for 2 min. Samples were rinsed 3 times for 5 min in 1X PBS and incubated for 1 h at 37°C with FITC-labeled dUTP in terminal deoxynucleotidyl transferase (TdT) buffer (30 mM Tris – HCl pH 7.2, 140 mm sodium cacodylate and 1 mm cobalt chloride) and TdT according to the protocol provided by the manufacturer (Roche). Positive controls were treated with DNase I for 15 min, and negative controls included sections incubated with FITC-labeled dUTP in the absence of TdT. Cells were washed in 1X PBS, mounted in Dako fluorescent mounting media (Dako S3023) and analyzed with a Zeiss Axiovert 200 M epifluorescence microscope equipped with an AxioCam HRm digital camera and AxioVision 4.6 software (Zeiss).

Hematoxylin and Eosin staining

Whole cervical, thoracic and lumbar DRGs were extracted from P15 mice and fixed in formalin (Fisher Scientific) at 4°C for 48 h. Samples were then transferred to 70% ethanol and stored at 4°C until processing. DRG samples were processed at the University of Ottawa (Department of Pathology and Laboratory Medicine), where they were set in agarose before embedding in paraffin wax with a LOGOS microwave hybrid tissue processor. Paraffin-embedded samples were cut by microtome at a thickness of 4 μm and stained for hematoxylin and eosin using a Leica ST5010 Autostainer XL combined with Leica CV5030 Glass Coverslipper. Stained slides were scanned with a MIRAX MIDI digital slide scanner (Zeiss). Images were acquired using 3DHISTECH Panoramic Viewer 1.15.4 at 20× magnification.

RNA-isolation and RT-PCR analysis

RNA was isolated from P15 DRG, spinal cord and cortex tissues, as well as from primary sensory neuron cultures from P15 mice using the RNeasy® Mini Kit (Qiagen), as per the manufacturer’s protocol. Samples were reverse-transcribed with RT² First Strand cDNA synthesis kit (Qiagen) using 1 μg total RNA. For cDNA from P15 tissues, we then further diluted the preparation to a 1 in 5 dilution using RNase-free water (Qiagen). For each of the Dst transcripts, the RT-PCR reaction is made up of 10 μl SsoFast™ EvaGreen® Supermix (Bio-Rad), 5 μl diluted cDNA, 1 μl of 10 μM forward + reverse primer mix and 4 μl RNase-free water. The RT-PCR reaction for actin transcript includes 10 μl SsoFast™ EvaGreen® Supermix (Bio-Rad), 5 μl diluted cDNA, 0.2 μl of 10 μM forward + reverse primer mix and 4.8 μl RNase-free water. The cycling parameters are the same for each target: (1) denature at 95°C for 10 min; (2) denature at 95°C for 10 s; (3) anneal at 60°C for 30 s; and (4) repeat steps 2–3 for 30 cycles. The forward primers for the Dst isoforms are Dst-A1 forward CTA CAT GTA CGT GGA GGA GCA, Dst-A2 forward GAG GGC TGT GCT TCG GAT AG, Dst-A3 forward GTC TCC AAG GAT GCA CCT AGG GAT, Dst-A1/A2/A3 reverse CAT CGT TTG CAC CAA TGC C, β-actin (actb) forward CCG TCA GGC AGC TCA TAG CTC TTC and β-actin (actb) reverse CTG AAC CCT AAG GCC AAC CGT. Products were run in a 2% agarose gel containing RedSafe™ nucleic acid staining solution (Froggabio), and bands were visualized under UV light.

Quantitative RT-PCR

Dst-A1/-A2/-A3 primer sets are the same as indicated above. Expression of Dst transcripts was normalized to Ppia and Rps18 (Bio-Rad PrimePCR™ pre-optimized primers) by ∆∆Ct method. These two gene products were found to have the lowest M value out of a series of different targets tested. Reaction mixtures for these are the same as for Dst as described above. Cycling parameters are the same for each target: (1) denature at 95°C for 10 min; (2) denature at 95°C for 10 s; (3) anneal at 60°C for 30 s; and (4) repeat steps 2–3 for 40 cycles. Runs were amplified using a Bio-Rad CFX96. Each genotype and treatment condition (Dst shRNA or scrambled control) was run as three biological replicates (except where indicated), and all samples were run in technical triplicate, with only those within a range of 0.2 Cq used for analysis.

Transmission electron microscopy

Mice were anesthetized by intraperitoneal injection of tribromomethanol (Avertin) and transcardially perfused with 5 mL of 1X PBS followed by 10 mL of Karnovsky’s fixative (4% paraformaldehyde, 2% glutaraldehyde [Sigma-Aldrich] and 0.1 M sodium cacodylate buffer [Electron Microscopy Sciences] in 1X PBS). Dorsal root axons were then extracted and separated according to spinal level (cervical, thoracic and lumbar) and fixed overnight at 4°C in the same fixative. Following fixation, each root was cut into a straight 1 mm segment. Segments were subsequently washed in 0.1 M sodium cacodylate buffer (Electron Microscopy Sciences) for 1 h at room temperature, and again overnight at room temperature. Segments were post-fixed with 1% osmium tetroxide (Electron Microscopy Sciences) and 1.5% potassium ferrocyanide trihydrate (for myelin staining; Acros Organics) in 0.1 M sodium cacodylate buffer for 1 h at room temperature. Samples were then washed 3 times in 0.2 M sodium cacodylate buffer, pH 7.4, for 15 min each. A second fixation step was performed in 2% glularaldahyde and 0.2 M sodium cacodylate buffer for 1.5 h at room temperature. These were washed 3 times in distilled water for 10 min and then dehydrated twice for 20 min for each step in a graded series of ethanol (Commercial Alcohols) from 30–50–70–85–95% ethanol in water. This was followed by two 30 min washes in 100% ethanol, two 15 min washes in 50% ethanol/50% acetone and two 15 min washes in 100% acetone (Fisher Scientific). Dorsal root axon segments were infiltrated in 30% spurr resin/acetone for 20 min, and once overnight, then in 50% spurr resin/acetone for 6 h and finally in 100% spurr resin (Electron Microscopy Sciences) overnight. Segments remained in 100% spurr resin, which was changed twice a day for 3 days at room temperature. All infiltration steps were performed on a rotator at low speed. Segments were embedded in fresh liquid spurr resin, oriented longitudinally inside the molds and then polymerized overnight at 70°C. Ultrathin sections (80 nm) were then collected onto 200 mesh copper grids and stained with 2% aqueous uranyl acetate (Electron Microscopy Sciences) and Reynold’s lead citrate (sodium citrate [Electron Microscopy Sciences] and lead nitrate [Electron Microscopy Sciences]). Sections were observed under a transmission electron microscope (Hitachi 7100) at 40 000×, 70 000× and 100 000× magnifications. An n = 2–3 was used for all P15 dorsal root axon samples, while an n = 1 was used for P20 and P40 dorsal root axon samples. Approximately 50 micrographs were examined at the cervical, thoracic and lumbar levels for each n.

Horizontal ladder test

The testing apparatus was composed of a 4-sided clear box with a Sony Handycam HDR-CX210 5.3 Megapixel Camcorder placed inside, facing upwards toward a horizontal ladder placed lengthwise along the edges of the box. Pup testing began at P10, with testing occurring every second day until the death of the Dst^dt mice. Each pup was placed in the middle of the ladder (rungs spaced 0.5 cm apart) and movements were video recorded throughout a 2 min testing period. Analysis of videos included counting total number of steps by forelimbs and by hindlimbs, as well as the total number of faults (paw falling through the rungs) per forelimb or hindlimb. The data presented is the number of faults normalized to the total number of steps taken (by forelimb or hindlimb). Statistical comparisons were made by Student’s t-test for each time point between the following pairs: Dst^dt-27J and Dst^dt-Tg4, Dst^dt-27J and their WT littermates and Dst^dt-Tg4 and their WT littermates.

Statistical analysis

Prism 5.0 GraphPad software was used for all statistical analysis, with the exception of quantitative RT-PCR data. For this we used either CFX software (Bio-Rad) for two-way comparisons, and qbase+3.0 software (Biogazelle) for multiple comparison tests. All two-way comparisons were done by two-tailed Student’s t-test. Multiple comparison tests were done by one-way Analysis of variance (ANOVA), followed by Tukey’s post hoc test. P-value < 0.05 was considered statistically significant (shown as ^*), while P-value < 0.01 is considered highly statistically significant (shown as ^**).

Supplementary Material

Supplementary Material is available at HMG online.

Acknowledgements

We are grateful to the Kothary laboratory for helpful discussions and to Dr Robin Parks’ laboratory for generously donating AdV-mCherry for use in our optimization experiments. We thank Samantha F Kornfeld for critical reading of the manuscript. A.L.G. is supported by an Ontario Graduate Scholarship.

Conflict of Interest statement: None declared.

Funding

Canadian Institute of Health Research (MOP-126085 to R.K.).